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Genetics. Apr 2010; 184(4): 985–997.
PMCID: PMC2865932

Alterations in DNA Replication and Histone Levels Promote Histone Gene Amplification in Saccharomyces cerevisiae


Gene amplification, a process that increases the copy number of a gene or a genomic region to two or more, is utilized by many organisms in response to environmental stress or decreased levels of a gene product. Our previous studies in Saccharomyces cerevisiae identified the amplification of a histone H2A-H2B gene pair, HTA2-HTB2, in response to the deletion of the other H2A-H2B gene pair, HTA1-HTB1. This amplification arises from a recombination event between two flanking Ty1 elements to form a new, stable circular chromosome and occurs at a frequency higher than has been observed for other Ty1-Ty1 recombination events. To understand the regulation of this amplification event, we screened the S. cerevisiae nonessential deletion set for mutations that alter the amplification frequency. Among the deletions that increase HTA2-HTB2 amplification frequency, we identified those that either decrease DNA replication fork progression (rrm3Δ, dpb3Δ, dpb4Δ, and clb5Δ) or that reduce histone H3-H4 levels (hht2-hhf2Δ). These two classes are related because reduced histone H3-H4 levels increase replication fork pauses, and impaired replication forks cause a reduction in histone levels. Consistent with our mutant screen, we found that the introduction of DNA replication stress by hydroxyurea induces the HTA2-HTB2 amplification event. Taken together, our results suggest that either reduced histone levels or slowed replication forks stimulate the HTA2-HTB2 amplification event, contributing to the restoration of normal chromatin structure.

CHANGES in gene copy number can cause significant phenotypic consequences. Gene amplification, a process that increases gene copy number to two or more, has been shown to occur as part of normal developmental processes, such as chorion gene amplification during oogenesis in Drosophila melanogaster (Spradling 1981; Orr et al. 1984; Claycomb et al. 2004; Tower 2004). In other cases, gene amplification has been shown to enhance growth under particular conditions; for example, amplification of a weakened lac operon in Escherichia coli allows improved growth on medium containing lactose as the carbon source (Hastings et al. 2004; Kugelberg et al. 2006). Recent studies have shown that changes in gene copy number are common in the human genome because copy number variants in humans occur genomewide, are polymorphic, and are sometimes associated with particular traits (McCarroll and Altshuler 2007). Moreover, gene amplification in humans is associated with some cancers (Stark and Wahl 1984; Hahn 1993; Graux et al. 2004; Vogt et al. 2004; Albertson 2006; Fridlyand et al. 2006).

One mechanism by which copy number changes occur is via recombination between repeated elements, such as transposable elements. In the yeast Saccharomyces cerevisiae, Ty retrotransposons have been shown to play a significant role in mediating genomic rearrangements, including gene amplifications (see Mieczkowski et al. 2006 for a review). These changes have been shown to occur under many different growth conditions, including selections for increased gene expression or drug resistance, after stress, or spontaneously (for examples, see Liebman et al. 1981; Brown et al. 1998; Rachidi et al. 1999; Hughes et al. 2000; Dunham et al. 2002; Koszul et al. 2004; Schacherer et al. 2004; Selmecki et al. 2006; Argueso et al. 2008). A recent study showed that Ty-mediated amplifications are a common occurrence when S. cerevisiae is grown under conditions limiting for glucose, phosphate, or sulfate (Gresham et al. 2008).

Several studies in S. cerevisiae have provided strong evidence that compromised DNA replication can increase the level of mitotic recombination and of Ty-mediated rearrangements. Cha and Kleckner (2002) showed that double-strand breaks in DNA occur at a greater frequency when replication is impaired by a mec1-ts mutation, thereby leading to increased mitotic recombination. Moreover, Vernon et al. (2008) demonstrated that in mec1 tel1 strains, the breakpoints of interchromasomal genomic rearrangements are associated with Ty elements. In another study, reduced levels of the replicative DNA polymerase α were shown to slow DNA replication and to cause greatly elevated frequencies of rearrangements, with many occurring by recombination between Ty elements (Lemoine et al. 2005). Finally, in a recent study, segmental duplications, many Ty-mediated, were shown to occur in a DNA-replication-dependent fashion (Payen et al. 2008). Schizosaccharomyces pombe appears to be similar to S. cerevisiae because elevated levels of mitotic recombination were shown to occur when replication forks were blocked artificially (Lambert et al. 2005). Taken together, these studies suggest that impaired replication forks are prone to suffer double-strand breaks, which in turn promote mitotic recombination.

A tight connection has also been established between histone levels, impaired DNA replication, and genome stability. In S. cerevisiae, early studies established that histone levels are correlated with DNA synthesis (Hereford et al. 1981; Osley and Hereford 1982). Furthermore, altered histone levels have been shown to cause problems with chromatin structure, chromosome segregation, transcription, and other aspects of cell growth (Meeks-Wagner and Hartwell 1986; Norris and Osley 1987; Clark-Adams et al. 1988; Norris et al. 1988). More recent evidence suggests that reduced histone levels in S. cerevisiae cause problems with DNA replication because repression of histone H4 gene transcription results in the elevation of replication-stress-induced transcripts, thereby suggesting that H4 depletion causes an impairment of fork movement (Wyrick et al. 1999). Consistent with this finding, another study showed that decreased histone H4 levels increases stalled replication forks, thereby increasing the frequency of homologous recombination (Prado and Aguilera 2005). Conversely, treatment of yeast cells with agents that damage DNA or that stall or slow replication forks decreases histone mRNA levels in both yeast and mammalian cells (Lycan et al. 1987; Gasch et al. 2001; Su et al. 2004), likely reflecting the effects of these agents on progression through the cell cycle. Overall, these studies have highlighted the importance of controlling histone levels to maintain genome stability (Gunjan et al. 2005).

Our previous work showed that one mechanism by which S. cerevisiae maintains proper histone levels is by amplification of a histone H2A-H2B locus, HTA2-HTB2 (Libuda and Winston 2006). This amplification was shown to occur in response to reduced H2A-H2B levels caused by deletion of the other H2A-H2B locus, HTA1-HTB1. Our work demonstrated that HTA2-HTB2 amplification occurs by formation of a small circular chromosome, the result of mitotic recombination between two Ty1 elements that flank a 40-kb region of chromosome II that contains HTA2-HTB2, CEN2, and three origins of replication. Furthermore, the amplification occurs at a much higher frequency than other Ty-Ty recombination events. To understand circumstances in which HTA2-HTB2 amplification might normally occur, we have performed a genetic screen to identify regulators of the HTA2-HTB2 amplification event. Among the genes identified in our screen, we identified factors required for the normal progression of DNA replication forks. On the basis of this finding, we also show that treating cells with hydroxyurea (HU) stimulates HTA2-HTB2 amplification. In addition, our mutant screen shows that alterations in histone stoichiometry stimulate the amplification event. Finally, our analysis has shown that a greatly reduced expression of the genes encoding histones H3 and H4 significantly increases the level of DNA replication-stress-induced transcripts. On the basis of these results, we propose a model in which environmental conditions that reduce histone levels, and hence impair replication fork movement, enhance the formation of the HTA2-HTB2 amplification to aid in the recovery and restoration of normal chromatin structure.


S.cerevisiae strains and media:

All S. cerevisiae strains used in this study (Table 1) are isogenic with a GAL2+ derivative of S288C (Winston et al. 1995). Standard methods for mating, sporulation, transformation, and tetrad analysis were used, and all media were prepared as described previously (Rose et al. 1990). Strains were constructed by either crosses or transformations and were grown at 30°. Gene deletions made as part of this study removed the following sequences (coordinates given from 5′ of the ATG to 3′ of the termination codon): rrm3Δ, 12 bp 5′ to 152 bp 3′; dpb3Δ, 12 bp 5′ to 34 bp 3′; dpb4Δ, 102 bp 5′ to 1 bp 3′; clb5Δ, 305 bp 5′ to 418 bp 3′; dun1Δ, 40 bp 5′ to 40 bp 3′; mec1Δ, 1 bp to 6884 bp of the ORF; and sml1Δ, 39 bp 5′ to 71 bp 3′. Sequences of oligonucleotides used are in the supporting information, Table S1. The (hht1-hhf1)Δ mutation removes the HHT1-HHF1 locus from 100 bp 3′ of the HHT1 ORF to 250 bp 3′ of the HHF1 ORF. The (hht2-hhf2)Δ mutation removes the HHT2-HHF2 locus from 26 bp 3′ of the HHT2 ORF to 18 bp 3′ of the HHF2 ORF. The coq1::HIS3 reporter was integrated by transformation with Bsu361-digested pDL1 (Libuda and Winston 2006). FY2784 was created by transforming FY1990 with pDM10, selection on 5-fluoroorotic acid (5-FOA) medium to lose pDM1, transformation with pMR102, and selection on 5-fluoroanthranilic acid media with 2% galactose to lose pDM10.

S. cerevisiae strains used in these studies


Plasmids were prepared from E. coli strains DH5α or MH1 according to standard procedures. The plasmids B155 (Libuda and Winston 2006), pSAB6 (Hirschhorn et al. 1995), pDL1 (Libuda and Winston 2006), pDL2 (Libuda and Winston 2006), pMR102 (Mann and Grunstein 1992), pHB59 (Hirschhorn et al. 1992), pCC64 (Clark-Adams et al. 1988), and pCC65 (Clark-Adams et al. 1988) have been described previously. pDM1 was constructed by amplifying the HHT2-HHF2 locus from plasmid pCC65 (Clark-Adams et al. 1988) and ligating it into pRS416 (Brachmann et al. 1998) after digestion with HindIII and BamHI. pDM10 was constructed by amplifying the HHT1-HHF1 locus from plasmid pCC64 (Clark-Adams et al. 1988) and ligating it into pRS414 (Brachmann et al. 1998) after digestion with HindIII and BamHI.

A screen to identify mutations that increase or decrease the frequency of HTA2-HTB2 amplification:

To identify mutations that alter the frequency of HTA2-HTB2 amplification, we relied on the observation that amplification of HTA2-HTB2 suppresses the inviability caused by (hta1-htb1)Δ (Libuda and Winston 2006). First, we crossed the S. cerevisiae nonessential gene deletion set (Giaever et al. 2002) by strain FY2789, which contains both (hta1-htb1)Δ and plasmid pSAB6 (with HTA1-HTB1). After selecting for diploids, we replica plated strains to 5-FOA to lose pSAB6 and then sporulated the strains as described previously (Giaever et al. 2002). To assess the level of viable progeny when (hta1-htb1)Δ was combined with each deletion, progeny were replica plated to select for haploids (CanR), the (hta1-htb1)Δ mutation (Leu+), and the deletion set mutation (G418R). From this primary screen, a large number of gene deletions were found to either increase or decrease the level of (hta1-htb1)Δ papillae relative to wild type. These candidates were then retested by repeating the screen for the subset of deletion set strains identified in the primary screen.

Hydroxyurea and methyl methanesulfonate treatments:

HU (Sigma-Aldrich) was dissolved in water and filter sterilized before adding to liquid YPD for a final concentration of 50, 100, or 200 mm. Methyl methanesulfonate (MMS) (Sigma-Aldrich) was dissolved in water and filter sterilized before adding to liquid YPD for a final concentration of 0.02%. Each solution was made fresh for each experiment. For the assays measuring Ty1-Ty1 recombination and HTA2-HTB2 amplification frequencies, cells were initially grown to 1 × 108 cells/ml at 30° in SC-his to maintain the coq1::HIS3 reporter. YPD containing the different concentrations of HU or MMS were inoculated with the SC-his cultures to a final concentration of 1 × 106 cells/ml. Following growth for 24 hr at 30°, cells were plated as described above. For Northern analysis of cells treated with HU, cells were grown in YPD at 30° to 1–2 × 107 cells/ml. Then, HU was added for a final concentration of 200 mm. Total RNA was prepared at the time points shown in Figure 3 using methods described previously (Swanson et al. 1991).

Figure 3.
HU treatment decreases histone transcripts and increases replication-stress-induced transcripts. Wild-type strains carrying the coq1::HIS3 reporter, which assays for HTA2-HTB2 amplification formation, were grown in rich medium, and HU was added ...

Assays for measuring Ty1-Ty1 recombination and HTA2-HTB2 amplification formation:

The recombination frequency between two Ty1 elements was measured using Ty1-URA3-Ty1 recombination reporters at YDRWTy1-5 on chromosome IV and YPLWTy1-1 on chromosome XVI (Libuda and Winston 2006). HTA2-HTB2 amplification was measured using the coq1::HIS3 construct (Libuda and Winston 2006), which is based on a method adapted from Chan and Botstein (1993). To measure Ty-Ty recombination and HTA2-HTB2 amplification, strains containing both coq1::HIS3 and one of the Ty1-Ty1 recombination reporters (FY2763, FY2764, FY2765, and FY2766) were grown to 1 × 108 cells/ml at 30° in SC-his medium to maintain the coq1::HIS3 reporter. Cells were then shifted to either YPD or YPD containing either HU or MMS as described below. Following growth for 24 hr at 30°, cells were plated on (1) YPD to measure the number of total cells in the culture; (2) 5-FOA plates to measure the number of cells that had undergone Ty1-Ty1 recombination at the Ty1-URA3-Ty1 reporter; (3) YPglycerol to measure the number of cells that had undergone a recombination at the coq1::HIS3 reporter to restore the COQ1 gene; and (4) SC-his plates with glycerol as the carbon source (SC-his + glycerol) to measure the number of cells that either contained the HTA2-HTB2 amplification or were disomic for chromosome II. These plating assays were done for the following number of independent cultures for each treatment: no treatment, 40 cultures; 50 mm HU, 9 cultures; 100 mm HU, 12 cultures; 200 mm HU, 22 cultures; and 0.02% MMS, 9 cultures. Contour-clamped homogeneous electric field (CHEF) gel Southern analysis was performed for His+ Gly+ colonies to determine the frequency of HTA2-HTB2 amplification among His+ Gly+ colonies for each treatment. The number of His+ Gly+ isolates with the HTA2-HTB2 amplification over the total number tested was as follows: (1) no treatment, 28/61; (2) 50 mm HU, 10/12; (3) 100 mm HU, 8/10; (4) 200 mm HU, 33/40; and (5) 0.02% MMS, 23/30. Ty1-Ty1 recombination and HTA2-HTB2 amplification formation frequencies were calculated as described previously (Libuda and Winston 2006) but with the above frequencies for the HTA2-HTB2 amplification among His+ Gly+ colonies factored in. The average frequency (amplification events/total cells) for each treatment was as follows: (1) no treatment—4.2 × 10−5; (2) 50 mm HU—9.0 × 10−5; (3) 100 mm HU—3.4 × 10−4; (4) 200 mm HU—7.6 × 10−4; and (5) 0.02% MMS—1.4 × 10−4. The fold changes represent the average increase compared to the average for untreated cultures, shown with the standard error.

Northern hybridization analysis, CHEF electrophoresis, and Southern hybridization analysis:

Northern hybridization analysis was performed as previously described, with all probes generated by labeling of PCR products (Ausubel et al. 1991). All primers used for PCR are listed in Table S1. The TPI1 probe has been previously described (Hirschhorn et al. 1992). Other probes were as follows: RNR3, from 1229 to 1430 bp from the ATG; HTB1, from 162 bp 5′ to 61 bp 5′ of the HTB1 ORF; HTA2, from 20 bp 3′ to 134 bp 3′ of the ORF; HHF1, from +314 bp from the ATG to 175 bp 3′ of the ORF; HHT2, from 91 bp 5′ to 1 bp 5′ of the ORF; HHF2, from +301 bp from the ATG to 198 bp 3′ of the ORF; and ACT1, from 367 bp to 1015 bp from the ATG. The analysis of intact yeast chromosomes by CHEF gels followed by Southern hybridization analysis was performed as previously described (Libuda and Winston 2006). Probes for the amplified region [HTA2-HTB2 and 3′YBLWTy1-1 (amplification probe B)] were also prepared as previously described (Libuda and Winston 2006).

Controlling histone H3-H4 gene-pair transcription:

Strains FY2784 and FY4 were grown at 30° to 1–2 × 107 cells/ml in YPgal, YP + 2% raffinose (YPraf) + 0.05% galactose, YPraf + 0.01% galactose, or YPraf + 0.005% galactose. Total RNA was prepared from these cultures as described previously (Swanson et al. 1991).


A screen for deletions that alter the frequency of HTA2-HTB2 amplification:

Our previous results demonstrated that the histone H2A-H2B gene pair, HTA2-HTB2, can amplify by recombination between flanking Ty1 elements to form a small circular chromosome (Libuda and Winston 2006). The frequency of the amplification event becomes significantly elevated in a mutant background in which histone H2A and H2B levels are reduced by deletion of the other H2A-H2B locus, HTA1-HTB1. This observation raised the possibility that the HTA2-HTB2 amplification occurs in wild-type cells when there is a need to increase H2A-H2B levels. To investigate what circumstances or factors may regulate the HTA2-HTB2 amplification event, we screened for gene deletions that either increase or decrease its frequency. This screen, described in detail in materials and methods, took advantage of our previous finding that HTA2-HTB2 amplification is required for survival in an (hta1-htb1)Δ background. To conduct the screen, we constructed double mutants that contain both (hta1-htb1)Δ and a deletion of each of the 4815 nonessential genes from the S. cerevisiae deletion set and tested them for their level of survival, an indicator of the frequency of HTA2-HTB2 amplification.

From this screen, we identified 102 deletions that decrease or eliminate (hta1-htb1)Δ viability and 53 deletions that increase (hta1-htb1)Δ viability (Table S2). Among the deletions that decrease or eliminate (hta1-htb1)Δ viability, we identified genes encoding an RNA polymerase II component (RPB4), a protein that interacts with RNA polymerases (NNF2), histone-related factors (UAF30, KAP114, SPT10, and HDA1), and a component of ribonucleotide reductase (RNR4). Notably, no deletions in genes encoding recombination proteins were identified in our screen, which is likely due to redundancy among some recombination mechanisms and because some recombination mutants fail to sporulate, a requirement for our screen. Among the deletions that cause an increase in (hta1-htb1)Δ viability, we identified genes encoding Mediator coactivator complex components (MED1, NUT1, SRB2), the CDK submodule of Mediator (SRB8, SRB9, SRB10, SRB11), the Lsm1-7 complex (LSM1, LSM6, LSM7), histones H3 and H4 (HHF1, HHT1, HHT2), and the DNA replication components (DPB3, DPB4, RRM3, and CLB5). Deletion of the genes identified that encode Mediator components appear to derepress transcription of HTA2 in a cell-cycle-dependent manner (our unpublished results). Increased expression of HTA2-HTB2 might suppress the requirement for the HTA2-HTB2 amplification. Among the deletions identified, we have focused on two sets that increase viability: the genes involved in DNA replication and the genes encoding histones H3 and H4.

Deletion of genes involved in DNA replication increases HTA2-HTB2 amplification frequency:

Our screen for deletions that increase (hta1-htb1)Δ viability identified four genes that influence DNA replication: DPB3, DPB4, RRM3, and CLB5 (Table S2). DPB3 and DPB4 encode the two nonessential components of DNA polymerase epsilon (Pol epsilon) and have been implicated in replication fork progression (Araki et al. 1991; Dua et al. 1998; Ohya et al. 2000). In addition, previous studies have demonstrated roles for Dpb3 and Dpb4 in telomeric silencing (Tsubota et al. 2006) and have shown that Dpb4 is also a component of the Isw2 complex (Iida and Araki 2004; McConnell et al. 2004). RRM3 encodes a helicase that has been shown to be required for the normal progression of DNA replication forks (Ivessa et al. 2000, 2002; Azvolinsky et al. 2006, 2009). Interestingly, Rrm3 has also been shown to be associated with Pol2, the catalytic subunit of Pol epsilon (Azvolinsky et al. 2006). CLB5 encodes a cyclin that activates Cdc28 during S phase (Epstein and Cross 1992; Kuhne and Linder 1993; Schwob and Nasmyth 1993). In clb5Δ mutants, S phase is extended and approximately half of the origins fail to fire (Epstein and Cross 1992; Kuhne and Linder 1993; Schwob and Nasmyth 1993; Donaldson et al. 1998; McCune et al. 2008). To confirm the effect of these deletions on HTA2-HTB2 amplification, (hta1-htb1)Δ/HTA1-HTB1 diploids that also contained the relevant deletion were constructed and analyzed by a tetrad viability assay (Libuda and Winston 2006). Briefly, in this assay, an (hta1-htb1)Δ/HTA1-HTB1 diploid is sporulated, and tetrads are dissected to determine the frequency of viability of (hta1-htb1)Δ progeny. Our earlier studies showed that ~13% of the (hta1-htb1)Δ progeny are viable due to formation of the HTA2-HTB2 amplification, which most likely occurs post-meiotically (Libuda and Winston 2006). Our results demonstrate that, when dpb3Δ, dpb4Δ, rrm3Δ, or clb5Δ is present, the (hta1-htb1)Δ progeny have a significantly higher level of viability than those in a wild-type background (Table 2; Figure 1A). To determine if the viable (hta1-htb1)Δ progeny with these gene deletions contained the HTA2-HTB2 amplification, CHEF gel Southern blots were performed. In this method, the presence of the HTA2-HTB2 amplification is assayed by the presence of two bands that hybridize to an HTA2-HTB2 probe: chromosome II, the normal location of this locus, and the amplification (Libuda and Winston 2006). The results of this analysis showed that, indeed, most (hta1-htb1)Δ colonies with these deletions do contain the amplification (Table 2; Figure 1B). The small number that do not contain the amplification likely are disomic for chromosome II, another event that allows for (hta1-htb1)Δ survival (Libuda and Winston 2006). Taken together, these results suggest that deletion of these four genes increases the frequency of the HTA2-HTB2 amplification, possibly due to stalled replication forks or extended S phase.

Figure 1.
DNA replication gene deletions increase the frequency of HTA2-HTB2 amplification in (hta1-htb1)Δ strains. (A) Diploids that were HTA1-HTB1/(hta1-htb1)Δ and also homozygous for deletion of genes involved in DNA replication (DPB3, DPB4, ...
Effects of DNA replication and histone gene deletions on (hta1-htb1)Δ viability and HTA2-HTB2 amplification

Treatment with hydroxyurea induces amplification formation in wild-type cells:

Our finding that the HTA2-HTB2 amplification frequency is increased by mutations that impair DNA replication suggests that amplification might occur more frequently in wild-type cells when replication is impaired. Therefore, we measured the amplification frequency after treatment with MMS, a DNA-damaging agent, or HU, which slows DNA replication (Feng et al. 2006; Alvino et al. 2007). To measure the amplification frequency in wild-type cells, we adapted a previously described method that can detect two copies of the amplified region in a wild-type background, the coq1::HIS3 reporter (Figure 2A; Chan and Botstein 1993; Libuda and Winston 2006; see materials and methods for details). To control for general effects on Ty1-Ty1 recombination, we also measured recombination frequencies for two different Ty1-URA3-Ty1 recombination reporters (Libuda and Winston 2006).

Figure 2.
HU treatment increases the frequency of HTA2-HTB2 amplification in wild-type strains. (A) Diagram of the coq1::HIS3 ploidy marker for monitoring the HTA2-HTB2 amplification event. An integrating plasmid containing an internal COQ1 restriction fragment ...

After treatment with MMS, the frequency of HTA2-HTB2 amplification formation increased ~3.5-fold, while Ty-URA3-Ty recombination was not significantly increased (Figure 2B; materials and methods). Previous studies (Parket and Kupiec 1992) also showed that MMS treatment does not increase Ty-Ty recombination. To assay the effect of slowed DNA replication, we measured HTA2-HTB2 amplification frequency after growth in different levels of HU. Our results showed that the amplification frequency increased with increasing HU concentrations, up to an 18-fold stimulation in 200 mm HU (Figure 2B; materials and methods). Moreover, Ty1-URA3-Ty1 recombination frequency was at most only modestly affected by HU (Figure 2B; materials and methods). The significant stimulation in amplification frequency upon treatment with HU implicates a role for DNA replication fork progression in HTA2-HTB2 amplification.

Previous studies have shown that impaired replication or DNA damage results in reduced levels of mRNAs from histone genes and increased levels of mRNAs from replication-stress-induced genes, such as RNR3 (Lycan et al. 1987; Elledge and Davis 1990; Gasch et al. 2001; Su et al. 2004; Zhao 2004). To test whether these transcriptional changes occur in our amplification assays, we measured both histone gene and RNR3 mRNA levels from cells treated with 200 mm HU. Our results show that histone gene mRNAs begin to decline after 5 min in 200 mm HU, reaching their minimal level by 15–30 min and then recovering to near-wild-type levels by 240 min (Figure 3). In addition, our results show that RNR3 mRNA levels increase during the 200 mm HU treatment, peaking at 120 min (Figure 3). As shown later, this increase is similar to the increase in RNR3 mRNA levels observed in cells with significantly reduced histone H3-H4 gene-pair expression. Overall, these results support previous findings that demonstrated changes in both histone and replication-stress-induced mRNA levels upon HU treatment (Lycan et al. 1987; Elledge and Davis 1990; Gasch et al. 2001). Moreover, these findings connect conditions that cause HTA2-HTB2 amplification, altered DNA replication, and decreased histone gene transcription.

Replication checkpoint pathway components are not required for HTA2-HTB2 amplification formation in (hta1-htb1)Δ strains:

Replication fork pauses and slowed DNA replication, as is observed with HU treatment, activate the replication checkpoint to assist with recovery from the pauses (Elledge and Davis 1990; Lambert and Carr 2005). Because we observed increased HTA2-HTB2 amplification frequency with HU treatment, we wanted to determine if the replication checkpoint pathway is required for this amplification event. Since deletions of genes encoding replication checkpoint pathway components cause HU sensitivity, we were unable to use the coq1::HIS3 reporter assay to address this question. Instead, we used the tetrad viability assay, measuring the frequency of the survival of (hta1-htb1)Δ progeny in homozygous deletions of MEC1 and DUN1, two genes required for the checkpoint pathway. For mec1Δ, the diploids were also homozygous for sml1Δ, which is required to suppress the lethality of mec1Δ (Zhao et al. 1998). Our results show that mec1Δ sml1Δ, dun1Δ, and sml1Δ do not significantly alter the frequency of HTA2-HTB2 amplification (Table 2). On the basis of CHEF gel analysis (Figure S1), most of the viable (hta1-htb1)Δ strains in those backgrounds contained the HTA2-HTB2 amplification. These results suggest that the replication checkpoint pathway is not required for HTA2-HTB2 amplification.

Histone H3-H4 gene-pair deletions increase (hta1-htb1)Δ viability but have different effects on HTA2-HTB2 amplification formation:

In our screen for deletions that increase (hta1-htb1)Δ viability, we also identified the two histone H3 genes, HHT1 and HHT2, and a histone H4 gene, HHF1. The S. cerevisiae H3-H4 genes are encoded in two divergently transcribed gene pairs, HHT1-HHF1 and HHT2-HHF2. On the basis of the screen results, we tested a deletion of each gene pair by the tetrad viability assay. In these tests, (hta1-htb1)Δ/HTA1-HTB1 diploids were also homozygous for either (hht1-hhf1)Δ or (hht2-hhf2)Δ. Each diploid was sporulated, followed by tetrad dissection and analysis to determine the frequency of viable (hta1-htb1)Δ progeny. Our results show that the deletion of each of the H3-H4 gene pairs caused a large increase in the viability of (hta1-htb1)Δ progeny—to 89% in the (hht1-hhf1)Δ strain background and 79% in the (hht2-hhf2)Δ strain background (Table 2, Figure 4A). To determine if the viable (hta1-htb1)Δ (hht1-hhf1)Δ and (hta1-htb1)Δ (hht2-hhf2)Δ strains contained the HTA2-HTB2 amplification, we performed Southern analysis of CHEF gels (materials and methods). Surprisingly, only 27% of (hht1-hhf1)Δ (hta1-htb1)Δ spores contained the amplification (Table 2, Figure 4B), suggesting that the (hht1-hhf1)Δ deletion can suppress the inviability caused by (hta1-htb1)Δ in the absence of any amplification. In contrast, 84% of (hht2-hhf2)Δ (hta1-htb1)Δ spores contained the amplification, strongly suggesting that (hht2-hhf2)Δ increases HTA2-HTB2 amplification frequency.

Figure 4.
Histone H3-H4 gene-pair deletions have different effects on HTA2-HTB2 amplification formation. (A) Tetrad dissections were performed on HTA1-HTB1/(hta1-htb1)Δ diploids that were also homozygous for deletion of one of the histone H3-H4 gene pairs ...

To confirm these distinct genetic interactions between the deletions of the two different H3-H4 loci and (hta1-htb1)Δ, we constructed each class of double mutant by crosses and then tested for viability. In each cross, one parent contained (hta1-htb1)Δ and also contained a plasmid with HTA1-HTB1 and URA3 (plasmid pSAB6). This strain did not contain any amplification. The second parent contained a deletion of one of the two H3-H4 loci, either (hht1-hhf1)Δ or (hht2-hhf2)Δ, and also did not contain the amplification. The viability of the histone double mutants was then tested in the progeny by the ability to grow on medium containing 5-FOA, a compound that counterselects for URA3. The (hta1-htb1)Δ (hht1-hhf1)Δ double mutants were 5-FOA resistant (10/10 progeny from 12 complete tetrads), confirming that (hht1-hhf1)Δ suppresses (hta1-htb1)Δ inviability. In contrast, the (hta1-htb1)Δ (hht2-hhf2)Δ double mutants were 5-FOA sensitive (7/7 progeny from 11 complete tetrads). This result suggests that (hht2-hhf2)Δ does not significantly suppress (hta1-htb1)Δ inviability. Taken together, these results suggest that, surprisingly, (hht1-hhf1)Δ suppresses the need for the HTA2-HTB2 amplification, whereas (hhf2-hht2)Δ enhances its frequency. We note that previous studies showed that HHT1-HHF1 is expressed at a substantially lower level than HHT2-HHF2 (Cross and Smith 1988). These differences likely lead to the different phenotypes with respect to HTA2-HTB2 amplification (see discussion).

Evidence that reduced histone H3-H4 levels cause paused DNA replication forks:

Previous studies demonstrated that repressing histone H4 gene transcription results in the elevation of replication-stress-induced transcripts (Wyrick et al. 1999). To determine if a reduction of both H3-H4 gene transcripts can also confer a replication stress signal, we constructed a strain in which the only copies of genes encoding histones H3 and H4 are under the control of the GAL1-10 promoter (materials and methods). Then we measured the level of RNR3 mRNA under conditions where HHT2-HHF2 is expressed at low levels. Our results show that RNR3 mRNA levels are significantly induced when HHT2-HHF2 transcripts are reduced (Figure 5). These results are consistent with previous studies (Wyrick et al. 1999) that suggest that an alteration in histone stoichiometry can increase replication-pause-induced transcripts. The increase in RNR3 mRNA levels is similar to that observed upon HU treatment (compare Figure 5 to Figure 3). Interestingly, we also observed that decreasing HHT2-HHF2 mRNA levels caused a decreased level of HTA2 mRNA. Taken together, these findings indicate a potential relationship between decreased histone gene transcription, increased replication-stress-induced transcription, and HTA2-HTB2 amplification.

Figure 5.
Reduced histone H3-H4 gene-pair transcript levels increase replication-stress-induced transcripts. A wild-type strain and a strain deleted for both histone H3-H4 loci [(hht1-hhf1)Δ (hht2-hhf2)Δ] but containing a plasmid ...


Our previous studies showed that when one histone H2A-H2B locus, HTA1-HTB1, is deleted, the second H2A-H2B locus, HTA2-HTB2, amplifies as part of a small circular chromosome. In this article, we have presented new results that suggest that this amplification event is stimulated in response to either of two mutually dependent changes: paused/slowed DNA replication forks or reduced histone gene transcription. First, as part of a genetic screen to identify mutations that alter the frequency of HTA2-HTB2 amplification formation, we identified several genes that encode factors involved in DNA replication. A common phenotype caused by mutations in these genes is the increased pausing of replication forks, suggesting that stalled forks induce HTA2-HTB2 amplification. Second, we showed that treatment with HU, a chemical that slows DNA replication, also stimulates HTA2-HTB2 amplification, thereby strongly supporting the idea that alterations in DNA replication induce this event. Third, our mutant screen also led to the demonstration that deletion of the histone H3-H4 locus, HHT2-HHF2, increases HTA2-HTB2 amplification. Finally, we have confirmed and extended previous results from other labs that have shown that slowing DNA replication by HU treatment results in reduced histone mRNA levels and, conversely, that reduced histone gene transcription induces stalled or slowed replication forks. Taken together, our findings suggest that amplification of HTA2-HTB2 is part of a cellular response that occurs when environmental changes perturb the replication of chromatin and the proper stoichiometry of histones.

Previous studies have shown that recombination proteins are recruited to sites of stalled replication forks (Lambert et al. 2005; Bailis et al. 2008). Those findings, taken together with our results, suggest that stalled or slowed replication forks within the amplified region are part of the mechanism that forms the HTA2-HTB2 amplification. Our results also demonstrate that the possible use of stalled or slowed replication forks as part of the signal to form the HTA2-HTB2 amplification is independent of the replication checkpoint. Current results do not yet explain why the Ty1-Ty1 recombination event that forms the HTA2-HTB2 amplification occurs at a much greater frequency than other Ty1-Ty1 recombination events. One obvious possibility is that either the mutants that we tested or HU treatment causes a more severe impairment of replication fork progression in the region of the amplification than in other genomic regions. However, previous genomewide studies of HU-treated cells (Feng et al. 2006; Alvino et al. 2007), of an rrm3Δ mutant (Azvolinsky et al. 2009), and of a clb5Δ mutant (McCune et al. 2008) do not support this idea. Thus, some aspect of impaired DNA replication that is yet to be elucidated likely accounts for the greatly enhanced frequency of Ty1-Ty1 recombination that forms the HTA2-HTB2 amplification.

The data presented here also suggest that histone H3-H4 levels play a key role in determining the response of cells to the reduced H2A-H2B levels caused by (hta1-htb1)Δ. Deletion of one H3-H4 gene pair, (hht2-hhf2)Δ, stimulates HTA2-HTB2 amplification in (hta1-htb1)Δ strains; conversely, deletion of the other H3-H4 gene pair, (hht1-hhf1)Δ, alleviates the requirement for HTA2-HTB2 amplification in (hta1-htb1)Δ strains. These differences are likely caused by the distinct effects that the two deletions have on H3-H4 levels because previous studies have shown that the two H3-H4 loci are transcribed at different levels, with HHT2-HHF2 producing five- to sevenfold more mRNA than HHT1-HHF1 (Cross and Smith 1988). Thus, the viability of the (hht1-hhf1)Δ (hta1-htb1)Δ double mutant suggests that the H3-H4 level in this mutant allows approximately normal stoichiometry between H2A-H2B and H3-H4, although the absolute histone levels may be reduced in the (hht1-hhf1)Δ (hta1-htb1)Δ double mutant compared to wild type. In contrast, in the (hht1-hhf2)Δ (hta1-htb2)Δ double mutant, where H3-H4 levels would be lower, histone stoichiometry would be greatly skewed; this acute alteration in histone stoichiometry may signal for amplification of HTA2-HTB2, thereby accounting for the increase in HTA2-HTB2 amplification frequency observed in these double mutants. Since amplification of HTA2-HTB2 clearly does not restore normal histone stoichiometry when H3-H4 levels are reduced, why would amplification of HTA2-HTB2 be a response to reduced H3-H4 levels? The HTA2-HTB2 amplification may represent one of several responses to an altered histone stoichiometry.

Many studies have now shown that S. cerevisiae carefully controls histone levels by several distinct mechanisms, including both transcriptional control (Osley 1991) and protein stability (Gunjan and Verreault 2003; Gunjan et al. 2005; Singh et al. 2009). Our studies demonstrate that HTA2-HTB2 amplification is yet another method of dosage compensation for altered histone levels. Why are there so many mechanisms to control histone levels? Clearly, either decreased or increased histone levels have a profound effect on growth. At the extreme, altered levels result in inviability (Rykowski et al. 1981; Kolodrubetz et al. 1982; Smith and Stirling 1988), whereas more modest effects can greatly impair transcription, chromosome stability, and DNA replication (Meeks-Wagner and Hartwell 1986; Clark-Adams et al. 1988; Wyrick et al. 1999; Prado and Aguilera 2005). Given these threats, and that histone levels can likely be perturbed by a multitude of mechanisms, it seems prudent to have several possible responses to conditions that alter proper histone levels.

Our model suggests than an alteration in histone stoichiometry causes the induction of several response mechanisms, one of which is the formation of the HTA2-HTB2 amplification (Figure 6). In this model, either altered histone stoichiometry [(hta1-htb1)Δ or (hht2-hhf2)Δ] or alterations in DNA replication (dpb3Δ, dpb4Δ, rrm3Δ, clb5Δ, or HU treatment) stimulate the HTA2-HTB2 amplification event. Since lowered histone H3-H4 gene transcription induces transcripts characteristic of stalled or slowed replication forks, we propose that altered histone stoichiometry may promote a signaling cascade in which stalled replication forks induce the Ty1-Ty1 recombination event that forms the HTA2-HTB2 amplification. Alternatively, as observed with the HU treatment, slowed DNA replication may reduce histone levels and, in turn, stimulate the specific Ty1-Ty1 recombination event. In addition to HTA2-HTB2 amplification, altered histone stoichiometry can induce other pathways known to restore normal histone stoichiometry, such as changes in histone gene transcription or histone protein stability. Overall, this model illustrates how a population of cells utilizes several mechanisms as a general response to altered histone levels.

Figure 6.
Model for a response to altered histone levels in a population of wild-type cells. After encountering an environmental condition that alters histone levels, a population of cells can attempt to restore normal histone stoichiometry by stimulating the ...

The HTA2-HTB2 amplification, which is the product of a directed Ty1-Ty1 recombination event, represents part of a cellular response for adapting to environmental changes that alter histone stoichiometry. Several studies have demonstrated that Ty elements, in particular their recombination, can provide a rich source for both genetic variation and adaptive genomic changes in response to specific environmental stresses (Rosenberg 2001; Infante et al. 2003; Zeyl 2004; Mieczkowski et al. 2006; Roth et al. 2006). To the best of our knowledge, our results provide some the first evidence that cells can stimulate a specific Ty1-Ty1 recombination event in response to an environmental condition or stress. How a cell can enhance a particular Ty1-Ty1 recombination event is currently unknown. Future studies identifying other directed Ty1-Ty1 recombination events will reveal how the cell can direct manipulation of the genome via transposable elements for adaptation to specific environmental changes.


We thank the members of the Winston lab and Maitreya Dunham for insightful discussion and comments. This work was supported by National Institutes of Health grant GM32967 to F.W. and by a National Science Foundation Graduate Fellowship and an Albert J. Ryan Foundation Illick Fellowship to D.E.L.


Supporting information is available online at http://www.genetics.org/cgi/content/full/genetics.109.113662/DC1.


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